Protein Standards and Ladders Support — Troubleshooting

Here are some suggestions:

• Check the gel type/percentage of the gel that was used. Depending on the gel type and/or percentage, all the bands may not be seen. For example, the smallest bands of the protein standard may not resolve on a very low percentage gel whereas the higher molecular weight bands may not resolve on a high percentage gel.
• Check the expiration date on the protein standard. Expired lots may result in faded or missing bands due to protein degradation.
• Check the storage conditions for the protein standard. Improper storage conditions will compromise the stability of the proteins in the standard.
• Make sure that the protein standard was not heated/boiled prior to loading on the gel. Our protein standards are ready to load and we do not recommend heating/boiling them as this may cause degradation of proteins in the standard.

Here are some suggestions:

• Make sure that the correct amount of standard is loaded per lane. Loading too much protein can cause smearing and this is a problem especially with silver stained gels.
• Bands will not be as well resolved in low percentage gels. Try using a higher percentage gel.
• If the bands look smeary and non-distinct after a western transfer/detection, this may be due to the antibody being too concentrated. Follow the manufacturer’s recommended dilution or determine the optimal antibody concentration by dot-blotting.

• While loading, take care to make sure that there is no cross-contamination from adjacent sample lanes.
• Make sure that the correct amount of standard is loaded per lane. Loading too much protein can result in extra bands and this is a problem especially with silver stained gels.
• Improper storage of the standard or repeated freeze/thawing can result in protein degradation.

Pre-stained standards have a dye that is covalently bound to each protein that will result in the standard migrating differently in different buffer systems (i.e., different gels). As a result, using a pre-stained standard for molecular weight estimation will only give the apparent molecular weight of the protein. Pre-stained standards may be used for molecular weight approximation, confirming gel migration and estimating blotting efficiency but for accurate molecular weight estimation, an unstained standard should be used.

• Increase voltage, current or length of time for transfer
• SDS in the gel and in the SDS-protein complexes promotes elution of the protein from the gels but inhibits binding of the protein to membranes. This inhibition is higher for nitrocellulose than for PVDF. For proteins that are difficult to elute from the gel such as large molecular weight proteins, a small amount of SDS may be added to the transfer buffer to improve transfer. We recommend pre-equilibrating the gel in 2x Transfer buffer (without methanol) containing 0.02–0.04% SDS for 10 minutes before assembling the sandwich and then transferring using 1x transfer buffer containing 10% methanol and 0.01%SDS.
• Methanol removes the SDS from SDS-protein complexes and improves the binding of protein to the membrane, but has some negative effects on the gel itself, leading to a decrease in transfer efficiency. It may cause a reduction in pore size, precipitation of some proteins, and some basic proteins to become positively charged or neutral. Make sure that the methanol concentration in the transfer buffer is not more than 10–20% and that high-quality, analytical grade methanol is used.

• Decrease voltage, current or length of transfer time
• Make sure that the methanol concentration in the transfer buffer is proper; use a methanol concentration of 10–20% methanol removes the SDS from SDS-protein complexes and improves the binding of protein to the membrane.
• Make sure that the SDS concentration (if added) in the transfer buffer is proper, don’t use more than 0.02–0.04% SDS. Using too much SDS can prevent binding of proteins to the membrane.
• Check the pore size of the membrane and the size of the target protein. Proteins smaller than 10kDa will easily pass through a 0.45 μm pore size membrane. If proteins smaller than 10 kDa are of interest, it would be better to use a 0.2 μm pore size membrane.

The fading is most likely due to detergent in the western blocking/washing solutions that can remove some of the proteins from the membrane. The dye itself will not wash off of the proteins because it is covalently bound. We have found that smaller pore size membranes retain the proteins better during blocking and wash procedures, and hence recommend use of 0.2 μm instead of 0.45 μm membranes for best resolution and protein retention. After transfer, it is a good idea to circle the pre-stained bands with a pencil on the membrane, so band positions can be identified after blocking and processing.

Here are some suggestions:

• Verify that the detection reagents are working well. Optimize the antibody concentration to obtain best results.
• Make sure that the amount of standard loaded on the gel is correct.
• Optimize the transfer conditions (current, voltage, transfer time).
• Enzyme-conjugated primary antibodies may not bind efficiently with the proteins in the MagicMark XP Western Protein Standard. We recommend using unconjugated primary antibody, followed by the addition of enzyme-conjugated secondary antibody.
• Note: The Anti-myc-AP/HRP and Anti-V5-AP/HRP antibodies do not bind to MagicMark XP proteins.

Here are possible causes and solutions:

Here are possible causes and solutions:

Here are possible causes and solutions:

Here are possible causes and solutions:

Here are possible causes and solutions:

Here are possible causes and solutions:

Here are possible causes and solutions:

Here are possible causes and solutions:

Here are possible causes and solutions:

Here are possible causes and solutions:

This is likely due to the dye-front not run off the gel or removed from the blot. We recommend running the dye front off of the gel or removing it from the blot.

Here are possible causes and solutions: